The authors thank Kerstin Glensk for preparing the MTases M.BseCI and Set7/9 and gratefully acknowledge funding by the Excellence Initiative of the German Federal and State Governments and RWTH Aachen University. The authors are happy to provide 6BAz and SeAdoYn or other cofactor analogues for collaborative research.

1. General Instructions
<ol>
	<li>Store aziridine cofactor 6BAz (in DMSO) and protein MTase Set7/9 at -80 &#176;C and all other reagents including double-activated cofactor SeAdoYn and DNA MTase M.BseCI (in 50% glycerol) at -20 &#176;C.</li>
	<li>Determine the concentration of 6BAz and SeAdoYn via UV/Vis spectroscopy using the extinction coefficients &#949;269nm&#160;(6BAz) = 16,000 cm-1 M-1 and &#949;260nm (SeAdoYn) = 15,400 cm-1 M-1 in deionized water. Determine the concentration of MTases by the Bradford assay or, if the extinction coefficient is available, via direct absorption at 280 nm.</li>
	<li>Try to avoid creating bubbles by intensive pipetting or vortexing to prevent loss of enzyme activity. Instead, mix by gently pipetting up and down.</li>
	<li>When adding aziridine cofactors from stock solutions in DMSO make sure that final DMSO concentration in the assay is less than 5%. Always include 10 mM magnesium ions in the assay buffer to prevent non-specific reactions with DNA.</li>
	<li>When adding double activated cofactors from acidic stock solutions use small volumes (highly concentrated stock solutions) to avoid pH changes and make sure that the pH of the assay solution does not change significantly. Avoid thiols, e.g., &#946;-mercaptoethanol or dithiothreitol (DTT), in the assay buffer because they can interfere with the click reaction by complexation of the required copper ions.</li>
</ol>
2. One-step Labeling of DNA via Aziridine Cofactors
<ol>
	<li>Sequence-specific Methyltransferase-Induced Label ing (SMILing) of plasmid DNA with M.BseCI DNA MTase and aziridine cofactor 6BAz.
	<ol>
		<li>Thaw the cofactor solution at 20 &#176;C and prepare the reaction mixtures on ice.</li>
		<li>In addition to the assay perform a &#8220;cofactor&#8221; control, to visualize any non specific modifications, and an &#8220; enzyme&#8221; control, to make sure that the MTase preparation is free of the natural cofactor AdoMet.</li>
		<li>For the assay mix 2 &#181;l of 10x modification buffer (containing 100 mM Tris-HCl, 100 mM MgCl2, 20 mM &#946;-mercaptoethanol, pH 7.4), 2 &#181;l of pBR322 (0.5 &#181;g/&#181;l), 10 eq. M.BseCI per recognition sequence on the DNA (1 recognition sequence in pBR322) and the aziridine cofactor 6BAz to a final concentration of 60 &#181;M within a total volume of 20 &#181;l. Add cofactor and DNA MTase last. 
		NOTE: &#946;-Mercaptoethanol is toxic, corrosive and environmentally damaging.</li>
		<li>For the &#8220;cofactor&#8221; control add deionized water instead of M.BseCI and for the &#8220;enzyme&#8221; control add deionized water instead of 6BAz.</li>
		<li>Mix the solutions by gently pipetting up and down.</li>
		<li>Incubate the tubes at 55 &#176;C for 1 hr.</li>
		<li>Centrifuge briefly to collect all liquid at the bottom of the tubes.</li>
	</ol>
	</li>
	<li>Restriction-modification assay to verify DNA modification.
	<ol>
		<li>Prepare a solution by mixing 10 &#181;l 10x R.TaqI buffer (containing 100 mM Tris-HCl, 50 mM MgCl2, 1 M NaCl, 1 mg/ml bovine serum albumin, pH 8.0), 80 &#181;l deionized water and 3.3 &#181;l of the restriction endonuclease (REase) from Thermus aquaticus (R.TaqI, 10 U/&#181;l). Make sure to add the REase in the last step.</li>
		<li>To each tube from 2.1.7 add 2 &#181;l of 10x R.&#160;TaqI buffer and 28 &#181;l of the solution from above (2.2.1).</li>
		<li>Mix the solutions by gently pipetting up and down.</li>
		<li>Incubate the tubes at 65 &#176;C for 30 min.</li>
		<li>Centrifuge briefly to collect all liquid at the bottom of the tubes.</li>
	</ol>
	</li>
	<li>Electromobility shift assay (EMSA) with streptavidin to verify functional modification.
	<ol>
		<li>Remove 25 &#181;l from each tube (2.2.5) and add 2.4 &#181;l of a streptavidin solution (1 mM with respect to streptavidin monomer in streptavidin buffer containing 100 mM Na2HPO4, 100 mM NaCl, pH 7.5; 4 equivalents of total biotin). Add 2.4 &#181;l of streptavidin buffer to the remaining tubes.</li>
		<li>Incubate all tubes at 37 &#176;C for 1 hr.</li>
	</ol>
	</li>
	<li>Analysis via agarose gel electrophoresis.
	<ol>
		<li>Add 5 &#181;l of 6x loading buffer (0.25% bromophenol blue, 30% glycerol) to each tube.</li>
		<li>Mix the solutions gently.</li>
		<li>Load 10 &#181;l of each sample into the wells of an agarose gel (1% agarose in 0.5x TBE buffer containing 1x GelRed from a 10,000x stock solution).</li>
		<li>Run the gel in 0.5x TBE buffer with 80 V for approx. 1 hr.</li>
		<li>Visualize DNA bands on a UV table (312 nm) with a CCD camera equipped with a filter (540 &#177; 50 nm). 
		NOTE: UV light is damaging to eyes and skin.</li>
	</ol>
	</li>
</ol>
3. Two-step Protein Labeling via Double Activated Cofactors
<ol>
	<li>Methyltransferase-Directed Transfer of Activated Groups (mTAG) with Set7/9 and double-activated cofactor SeAdoYn for histone H3 lysine 4 labeling (modification step).
	<ol>
		<li>Thaw the components and prepare the reaction mixtures on ice. NOTE: Always keep SeAdoYn cooled to avoid degradation.</li>
		<li>In addition to the assay perform a &#8220;cofactor&#8221; control, to visualize any non specific modifications, and an &#8220;enzyme&#8221; control, to exclude non-specific reactions of the fluorescent probe.</li>
		<li>Prepare an assay solution (20 &#181;l) containing modification buffer (50 mM Tris-HCl, 5% glycerol, pH 8.5), 10 &#181;M histone H3, 10 &#181;M Set7/9 and 600 &#181;M SeAdoYn (mixture of both epimers at selenium). In the last steps add cofactor and then MTase.</li>
		<li>For the &#8220;cofactor&#8221; control prepare an assay solution as in 3.1.3 and add 60 mM AdoMet to compete with the synthetic cofactor. For the &#8220;enzyme&#8221; control add deionized water instead of SeAdoYn.</li>
		<li>Mix the solutions by slowly pipetting up and down. Check the pH by adding 1 &#181;l of each solution to the upper field of a pH strip (pH range 5 - 10).</li>
		<li>Incubate at 37 &#176;C for 2 hr.</li>
		<li>In the meanwhile prepare a 12% SDS polyacrylamide gel (running gel: 357 mM Bis-Tris pH 6.5-6.8, 0.1% (w/v) APS, 0.04% (v/v) TEMED and 12% acrylamide/bisacrylamide 37.5:1; loading gel: 357 mM Bis-Tris pH 6.5-6.8, 0.1% (w/v) APS, 0.04% (v/v) TEMED and 5% acrylamide/bisacrylamide 37.5:1). 
		NOTE: Acrylamide/bisacrylamide is toxic and health hazardous. Wear gloves during this procedure.</li>
	</ol>
	</li>
	<li>Chemical labeling of alkinylated lysine 4 in histone H3 via copper-catalyzed azide-alkyne cycloaddition (CuAAC) (labeling step).
	<ol>
		<li>Just before the end of the modification reaction prepare a 5x click mix containing 3 mM CuSO4, 3 mM tris(3-hydroxypropyl-triazolylmethyl)amine (THPTA), 250 mM sodium ascorbate and 6 mM TAMRA azide with a total volume of 20 &#181;l.</li>
		<li>Add 5 &#181;l of the freshly prepared 5x click mix to each tube to start the CuAAC and quench the modification reaction.</li>
		<li>Mix gently by pipetting up and down.</li>
		<li>Protect all tubes with aluminum foil from light to avoid photo-bleaching of the fluorophore.</li>
		<li>Incubate at 20 &#176;C for 1 hr.</li>
	</ol>
	</li>
	<li>Protein precipitation to remove excess of free TAMRA fluorophore.
	<ol>
		<li>To avoid outshining of the fluorescent labeled histone H3 by intensive in-gel fluorescence of free TAMRA fluorophore, remove excess fluorophore by precipitation of proteins (3.3.2 &#8211; 3.3.4)34.</li>
		<li>Add 75 &#181;l methanol, 18.8 &#181;l chloroform and 50 &#181;l deionized water to each tube and vortex briefly after each addition. Centrifuge at 16,000 x g for 5 min. Remove the upper phase without disturbing the interface layer, which contains the protein.</li>
		<li>Add 56.3 &#181;l methanol to the remaining phase in each tube, vortex and centrifuge at 16,000 x g for 5 min to pellet the protein. Remove the supernatant. Repeat this step to wash the pellet.</li>
		<li>Cover the open tubes with a lint free tissue and let them dry for 15 &#8211; 30 min.</li>
	</ol>
	</li>
	<li>Analysis via SDS PAGE.
	<ol>
		<li>Dissolve the precipitated proteins from 3.3.4 in 20 &#181;l SDS loading buffer (50 mM Tris-HCl, 2.5% (w/v) SDS, 10% (v/v) glycerol, 320 mM &#946;-mercaptoethanol and 0.05% (w/v) bromophenol blue, pH 6.8). Make sure to completely dissolve the pellet by rinsing the walls of the tubes with a pipette.</li>
		<li>Incubate the samples at 95 &#176;C for 10 min and let them cool down to 20 &#176;C.</li>
		<li>Centrifuge briefly to collect all liquid at the bottom of the tubes.</li>
		<li>Load the whole amount of each sample into the wells of an SDS polyacrylamide gel (3.1.7). Use 50 mM MOPS, 50 mM Tris-X (Tris-base), 5 mM EDTA, 0.1% (w/v) SDS as running buffer for electrophoresis.</li>
		<li>Run the gel with 120 V for approx. 90 min.</li>
		<li>Visualize the in-gel fluorescence on a UV table (312 nm) with a CCD camera equipped with a filter (540 nm &#177; 50 nm). 
		NOTE: UV light is damaging to eyes and skin.</li>
	</ol>
	</li>
</ol>

The authors disclose the following competing financial interest: E.W. is inventor on related patents.

One-step labeling of DNA with DNA MTases and aziridine cofactors (SMILing DNA) is a robust method but some aspects should be considered when planning the experiment.
Aziridine cofactor: The 6BAz concentration for DNA labeling with M.BseCI was 60 &#181;M. When using other DNA MTases the cofactor concentration should be optimized, e.g. concentrations as low as 20 &#181;M have been employed with the DNA MTase M.TaqI19. Low 6BAz concentrations have the advantage that a fourfold excess of streptavidin (binding sites with respect to the whole amount of biotin in the assay) can be directly added after incubation with REase without interfering with analysis by EMSA. Otherwise the biotinylated DNA should be purified to remove excess cofactor and avoid smearing of the DNA during agarose gel electrophoresis. When adding aziridine cofactors from stock solutions in DMSO make sure that the final DMSO concentration in the assay is less than 5%. Too much DMSO can inactivate enzymes. Always store aziridine cofactors at -80 &#176;C to avoid degradation.
DNA MTase: Enzymatic coupling of aziridine cofactors with DNA can lead to strong product inhibitors and the DNA MTases have to be used in stoichiometric amounts. Therefore always use more than 1 equivalent of DNA MTase. Most DNA MTases copurify with bound AdoMet which needs to be removed by extensive dialysis or washing the enzymes on a column with large volumes of buffer. The &#8220;enzyme&#8221; control will tell whether AdoMet is still present in the enzyme preparation. Labeling with M.BseCI can also be done by incubation over night at 37 &#176;C.
Modification buffer: Include 10 mM magnesium ions in the buffer to prevent non-specific reactions of aziridine cofactors with DNA. If magnesium ions lead to activation of contaminating nucleases, barium ions might be added instead.
For the sequence specific labeling of DNA not only SMILing DNA but also mTAG can be employed. Here double activated cofactors are used for enzymatic transfer of side chains with unique functional groups to DNA. These functional groups, e.g. primary amines, can be modified with a wide variety of reporter groups, including biotin or fluorophores, by NHS ester chemistry in a second step8,35,36. Alternatively we have started to synthesize double activated AdoMet analogues with large side chains already containing the reporter group and used them for DNA labeling in one step.
The two-step mTAG labeling procedure for proteins with the double activated cofactor SeAdoYn has been successfully used with a number of protein MTases30,31. However, the following comments should be considered before performing the experiment.
Double activated cofactor: These cofactors, including SeAdoYn, are stored under acidic conditions and care must be taken that the pH of the modification solution does not significantly change upon cofactor addition. We always check the pH by adding 1 &#181;l of the modification solution to a pH strip and, if the pH is too low, add small amounts of sodium hydroxide solution (50 mM) to adjust the pH. In addition cofactor analogues should be added in a low volume to avoid pH changes. Thus, minimal cofactor concentrations needed for full modification should be determined for other MTases and highly concentrated cofactor stock solutions are preferred. Cofactor concentrations are typically varied between 10 &#181;M and 600 &#181;M. The chemical synthesis of double-activated cofactors typically yields an approximately 1:1 mixture of epimers at sulfur or selenium and separation of the biologically active epimer, corresponding to the S-epimer of AdoMet, from the inactive epimer is often difficult to achieve. Thus, cofactor concentrations are typically given for a mixture of isomers. Double activated cofactor analogues are quite unstable at room temperature and should be stored at -20 &#176;C. Also make sure to thaw the stock solutions on ice and keep them cold during pipetting. They can be frozen again but to ensure reproducible activity we recommend to make aliquots.
Protein MTase: For transformations with double activated cofactors the MTases can be used in catalytic amounts. However, MTases are often slow enzymes having turnover numbers in the min-1 range with the natural cofactor AdoMet. For practical reasons we often use the MTases in stoichiometric amounts and minimize incubation time and temperature (typically 10 min to 2 hr and 20 &#176;C to 37 &#176;C). The &#8220;cofactor&#8221; control varies, depending on the type of reporter group and detection. For biotin labeling the MTase is simply omitted and analysis can be done by Western blotting (Figure 5B). For fluorescence labeling with in-gel-fluorescence detection the free fluorophore should be removed from the labeled protein before electrophoresis. In the current protocol this is achieved by precipitation of histone H3 in the presence of Set7/9 MTase. However, without Set7/9 precipitation is incomplete. Therefore, the &#8220;cofactor&#8221; control is performed in the presence of protein MTase but a high concentration of the natural cofactor AdoMet is added to displace SeAdoYn from the active site and prevent enzymatic transfer of the propargyl group (Figure 5A).&#160;However, if precipitation of histone H3 leads to loss of protein, SDS-PAGE can be extended and excess of free TAMRA fluorophore will run out of the gel with the&#160;bromphenol blue front.
Modification buffer: The optimal enzyme buffer can be used, but thiols, like dithiothreitol (DTT) and &#946;-mercaptoethanol, should be avoided. They bind to copper ions and block the following CuAAC click reaction.
CuAAC click reaction: The final concentration of TAMRA azide should be twice as high as the alkyne concentration. When using biological extracts the copper and ligand concentration should be increased up to 5 mM each. TAMRA azide is better soluble in DMSO than in water. Make sure that the final concentration of DMSO is less than 12%. Extended incubation times can lead to protein damage and smearing on the SDS polyacrylamide gel. Thus, the labeling reaction should be either stopped by protein precipitation or addition of thiols.
This two-step labeling procedure can also be used to label protein MTase substrates with biotin either for Western blot detection (Figure 5B) or isolation with streptavidin-coated magnetic beads. In addition, double activated AdoMet analogues can be used to label DNA, RNA and small natural products with corresponding MTases.
Compared to most other labeling methods for nucleic acids and proteins, MTase-mediated labeling has the advantage that native substrates can be directly used and no further modification is required. Of course, care must be taken that the natural substrates are not blocked by methylation through a corresponding MTase in vivo. In addition, MTase-mediated labeling is very flexible both in terms of reporter groups, which include biotin, fluorophores or other molecular tags, as well as targets for MTases. REBASE, a data base for restriction endonucleases and DNA MTases, lists more than 700 experimentally characterized DNA MTases with hundreds of different recognition sequences ranging from two to eight bp37 and it is expected that more than 200 different MTases of all types are produced in the human cell38. An important determinant for MTase activity is whether the AdoMet analogue fits into the enzyme active site. Although the presented cofactors 6BAz and SeAdoYn are designed to have small reactive groups, some MTases may not readily accept them. In these cases the active sites could be enlarged by protein engineering as has been demonstrated for DNA35, RNA23 and protein MTases25&#8211;28 with sterically more demanding double activated AdoMet analogues.
Sequence specific labeling of DNA and RNA with AdoMet analogues is of major interest for genotyping (DNA mapping)36 and methylation detection39, functional studies of DNA/RNA and DNA/RNA-modifying enzymes15 as well as for (nano)biotechnology13,14 and gene delivery19. Other methods for sequence specific covalent DNA labeling rely on synthetic triple helix-forming oligodeoxynucleotides, peptide nucleic acids or hairpin polyamides as targeting devices or on sequence specific nicking endonucleases (NEases) followed by nick translation labeling.1,2 However, the number of target sequences for triple helix formation or NEases (about 30 NEases are listed in REBASE37) are limited which makes DNA MTase-mediated labeling more general.
Specific labeling of proteins with double activated AdoMet analogues has been mainly directed towards applications in proteomic research, e.g. identification of new substrates for protein MTases in complex biological mixtures27,28, but other applications, like functional studies, should also be feasible. Although MTase-mediated labeling has been mainly performed with purified MTases in vitro, labeling can also be achieved in living cells as has been reported recently40. Of course, many other methods for specific protein labeling are available3,4. Besides incorporation of unnatural amino acids using engineered cells they typically require genetic fusions of the protein of interest with self-labeling tags/proteins or tags for enzyme-mediated labeling. In this respect short peptide sequences serving as substrates for protein MTases, e.g. N-terminal histone tails, could be fused to a protein of interest and specifically labeled with AdoMet analogues and corresponding protein MTases.
Finally, the biocatalytic repertoire of small molecule MTases can be expanded with synthetic AdoMet analogues41&#8211;44. This represents a new approach to introduce structural diversity into natural products, e.g. antibiotics or polyketides, which should find interesting applications in screening for novel biological activities.

Specific labeling of nucleic acids1,2 and proteins3,4 is of major interest for functional characterizations, medical diagnosis and (nano)biotechnology. Here we present an enzymatic labeling method for these biopolymers which is based on S-adenosyl-l-methionine (AdoMet or SAM)-dependent methyltransferases (MTases). This class of enzymes (EC 2.1.1.) targets individual nucleophilic positions (nitrogen, oxygen, sulfur and carbon atoms) within specific residues of nucleic acids and proteins and naturally transfers the activated methyl group of the cofactor AdoMet (Figure 1A)5. In addition, MTases can utilize synthetic cofactor analogues for specific labeling with affinity tags, fluorophores or other labels (Figure 1B)6. Two classes of AdoMet analogues have been developed: Aziridine cofactors for Sequence-specific Methyltransferase-Induced Labeling (SMILing)7 and double activated AdoMet analogues for methyltransferase-directed Transfer of Activated Groups (mTAG)8.
<img alt="Figure 1" fo:content-width="7in" src="/files/ftp_upload/52014/52014fig1highres.jpg" width="700" /> 
Figure 1: Reactions catalyzed by methyltransferases (MTases). A. Methyl group transfer from the natural cofactor AdoMet (SAM) to various substrates including DNA, RNA, proteins and small biomolecules. B. Labeling/functionalization of nucleic acids and proteins (NNNNN = base pairs for DNA, nucleotides for RNA and amino acids for proteins; XXXXX = recognition sequence of the MTase with target residue in green) with synthetic cofactor analogues. Aziridine cofactors containing a reporter group (blue sphere) attached to the adenine ring are sequence specifically coupled with the target residue (left) and double-activated AdoMet analogues lead to transfer of extended alkyl chains carrying a chemical reporter Y (right) which can be labeled by bioorthogonal click reaction in a second step. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52014/52014fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Aziridine cofactors work best with DNA MTases. They contain a three membered ring with a nitrogen atom9 (or an N-mustard10,11) instead of the sulfonium center as reactive group. Protonation of this nitrogen atom activates the aziridine ring for nucleophilic attack by the target nucleotide which leads to covalent coupling of the whole cofactor with DNA. By attaching reporter groups to the adenine ring the aziridine cofactors can be used in combination with DNA MTases to label DNA in one step (Figure 1B, left)7,12. This is demonstrated in detail for the biotinylation of DNA with 6BAz13&#8211;15 (aziridine cofactor with biotin attached to the 6 position of the adenine ring) and the adenine-specific DNA MTase from Bacillus stearothermophilus (M.BseCI)16 (Figure 2, see protocol section 2: One-step labeling of DNA via aziridine cofactors). In addition to M.BseCI (5&#8217;-ATCGAT-3&#8217; recognition sequence), the DNA MTases from Thermus aquaticus (M.TaqI, 5&#8217;-TCGA-3&#8217;), from Haemophilus heamolyticus (M.HhaI, 5&#8217;-GCGC-3&#8217;) and from Spiroplasma (M.SssI, 5&#8217;-CG-3&#8217;) have been successfully used to biotinylate DNA with 6BAz17. Furthermore, aziridine cofactors can be employed for one-step fluorescence DNA labeling18,19.
<img alt="Figure 2" fo:content-width="7in" src="/files/ftp_upload/52014/52014fig2highres.jpg" width="700" /> 
Figure 2: Sequence specific one-step biotinylation of DNA with M.BseCI and 6BAz. The DNA MTase M.BseCI recognizes the double-stranded DNA sequence 5&#8217;-ATCGAT-3&#8217; and naturally methylates the amino group of the second adenine residue (green) using AdoMet. With the aziridine cofactor 6BAz the course of the reaction is changed and M.BseCI leads to sequence specific DNA biotinylation by coupling the whole cofactor including biotin (blue) with the target adenine. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52014/52014fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Double activated AdoMet analogues contain extended unsaturated side chains instead of a methyl group at the sulfonium center (Figure 1B, right)20. The unsaturated double or triple bond in &#946;-position to the sulfonium center electronically compensates unfavorable steric effects within the transition state by conjugative stabilization. Since both the sulfonium center and the unsaturated bond activate the side chain for enzymatic transfer, these cofactors were named double-activated AdoMet analogues. Typically, they are used to transfer side chains with unique chemical groups (chemical reporters), like amino, alkyne and azide groups, for chemo-selective labeling in a second step8,21. In general, double-activated AdoMet analogues can not only function as cofactors for DNA MTases8,20,21 but also for RNA MTases22,23 and protein MTases24&#8211;28 allowing additional labeling of RNA and proteins. However, the extended side chains are sterically more demanding than a methyl group and enlarging the MTase active sites by protein engineering is often required to obtain efficient transfer rates. Another solution to this problem is to use an AdoMet analogue with a small propargyl group (three carbons) where the terminal alkyne serves two functions: 1. Stabilization of the transition state during enzymatic transfer and 2. reactive handle for following chemical modifications by copper-catalyzed azide-alkyne cycloaddition (CuAAC) click chemistry. It turned out that the resulting propargylic AdoMet analogue29 is quite unstable under neutral or slightly basic conditions and only of limited use. This drawback can be fixed by replacing the sulfur atom with selenium. The resulting cofactor 5&#8216;-[(Se)[(3S)-3-amino-3-carboxypropyl]prop-2-ynylselenonio]-5&#8216;-deoxyadenosine (SeAdoYn, Figure 3) is accepted by wild-type DNA, RNA and protein MTases30&#8211;32 which abrogate the need for protein engineering in many cases. This is exemplified by fluorescence protein labeling with the histone H3 lysine 4 (H3K4) MTase Set7/933 (Figure 3, see protocol section 3: Two-step protein labeling via double activated cofactors).
<img alt="Figure 3" fo:content-width="7in" src="/files/ftp_upload/52014/52014fig3highres.jpg" width="700" /> 
Figure 3: Sequence-specific two-step fluorescence labeling of histone H3 with Set7/9, SeAdoYn and TAMRA azide. The protein MTase Set7/9 naturally methylates the amino group of lysine 4 in histone H3 (H3K4, green) using AdoMet. With the double-activated cofactor SeAdoYn the MTase transfers a small propargyl group (red) to the lysine residue. The attached terminal triple bond is then selectively modified in a bioorthogonal click reaction (copper-catalyzed azide-alkyne cycloaddition, CuAAC) with azide-derivatized TAMRA (tetramethylrhodamine, blue) fluorophore. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52014/52014fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="0" cellpadding="0" cellspacing="0" width="669">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Name of Reagent/ Equipment</td>
			<td style="width:139px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">6BAz</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>Synthesized according to Weinhold et al., Patent number US 8,129,106, published March 6, 2012.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">&#946;-Mercaptoethanol</td>
			<td style="width:139px;">Serva</td>
			<td align="right" style="width:119px;">28625</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Acetic acid</td>
			<td style="width:139px;">Fisher Scientific</td>
			<td align="right" style="width:119px;">10304980</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Acrylamide/Bis Solution, 37.5:1</td>
			<td style="width:139px;">Serva</td>
			<td align="right" style="width:119px;">10688</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">UltraPure Agarose</td>
			<td style="width:139px;">Invitrogen</td>
			<td align="right" style="width:119px;">16500100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Ammonium persulfate (APS)</td>
			<td style="width:139px;">Serva</td>
			<td align="right" style="width:119px;">13375</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Bis-Tris</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">1304</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Boric acid</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">1115</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Bromophenol blue Na salt</td>
			<td style="width:139px;">Serva</td>
			<td align="right">15375</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Copper(II) sulfate</td>
			<td style="width:139px;">Aldrich</td>
			<td style="width:119px;">C1297</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Chloroform</td>
			<td style="width:139px;">Fisher Scientific</td>
			<td align="right">10020090</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Coomassie Brilliant Blue</td>
			<td style="width:139px;">Serva</td>
			<td align="right" style="width:119px;">17525</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">EDTA disodium salt</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">1034</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Ethanol</td>
			<td style="width:139px;">Merck</td>
			<td align="right" style="width:119px;">100983</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">GelRed (10000x in water)</td>
			<td style="width:139px;">Biotium</td>
			<td align="right" style="width:119px;">41003</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Glycerol (99.5%)</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">2006</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">FastRuler Low Range DNA Ladder</td>
			<td style="width:139px;">Thermo Scientific</td>
			<td style="width:119px;">SM1103</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Histone H3</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>Expressionplasmid obtained from Dr. Philipp Voigt and Prof. Danny Reinberg; expression and isolation according to T. J. Richmond et al., J. Mol. Biol. 1997, 272, 301-311.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">M.BseCI</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>Expressionplasmid obtained from Dr. Michael Kokkinidis; expression and isolation according to Kapetaniou et al., Acta Cryst. 2006, F63, 12-14.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Methanol</td>
			<td style="width:139px;">Fisher Scientific</td>
			<td align="right" style="width:119px;">10675112</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Magnesiumchloride&#160; hexahydrate</td>
			<td style="width:139px;">J.T. Baker</td>
			<td align="right" style="width:119px;">4003</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">MOPS</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">1081</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Sodium chloride</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">1112</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">pH strip (Neutralit)</td>
			<td style="width:139px;">Merck</td>
			<td align="right" style="width:119px;">1,095,330,001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">pBR322</td>
			<td style="width:139px;">Thermo Scientific</td>
			<td style="width:119px;">SD0041</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">R.TaqI (10u/&#181;l)</td>
			<td style="width:139px;">Thermo Scientific</td>
			<td>ER0671</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">SeAdoYn</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>Synthesized according to Willnow et al., ChemBioChem 2012, 13, 1167-1173.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Set7/9</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>Expressionplasmid obtained from Prof. Danny Reinberg, expression and isolation according to D. Reinberg et al., Genes Dev.2002, 16, 479-489.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Streptavidin</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">3058</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">(+)-Sodium L-ascorbate</td>
			<td style="width:139px;">Sigma Life Science</td>
			<td style="width:119px;">A7631</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">SDS Granular</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">1833</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">di-Sodiumhydrogenphosphat</td>
			<td style="width:139px;">Merck</td>
			<td align="right" style="width:119px;">106,586</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">TAMRA-azide</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">TaqI buffer (10x)</td>
			<td style="width:139px;">Thermo Scientific</td>
			<td style="width:119px;">B28</td>
			<td></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:245px;">N,N,N&#39;,N&#39;-Tetramethylethylenediamine (TEMED)</td>
			<td style="width:139px;">Acros Organics</td>
			<td align="right" style="width:119px;">42058</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Tris-HCl</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">1028</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Tris-X (TRIS-base)</td>
			<td style="width:139px;">Gerbu</td>
			<td align="right" style="width:119px;">1018</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:245px;">Tris(3-hydroxypropyltriazolyl-methyl)amine (THPTA)</td>
			<td style="width:139px;">Sigma-Aldrich</td>
			<td align="right" style="width:119px;">762342</td>
			<td></td>
		</tr>
	</tbody>
</table>

sequence-specific labeling of nucleic acids and proteins with methyltransferases and cofactor analogues

S-Adenosyl-l-methionine (AdoMet or SAM)-dependent methyltransferases (MTase) catalyze the transfer of the activated methyl group from AdoMet to specific positions in DNA, RNA, proteins and small biomolecules. This natural methylation reaction can be expanded to a wide variety of alkylation reactions using synthetic cofactor analogues. Replacement of the reactive sulfonium center of AdoMet with an aziridine ring leads to cofactors which can be coupled with DNA by various DNA MTases. These aziridine cofactors can be equipped with reporter groups at different positions of the adenine moiety and used for Sequence-specific Methyltransferase-Induced Labeling of DNA (SMILing DNA). As a typical example we give a protocol for biotinylation of pBR322 plasmid DNA at the 5&#8217;-ATCGAT-3&#8217; sequence with the DNA MTase M.BseCI and the aziridine cofactor 6BAz in one step. Extension of the activated methyl group with unsaturated alkyl groups results in another class of AdoMet analogues which are used for methyltransferase-directed Transfer of Activated Groups (mTAG). Since the extended side chains are activated by the sulfonium center and the unsaturated bond, these cofactors are called double-activated AdoMet analogues. These analogues not only function as cofactors for DNA MTases, like the aziridine cofactors, but also for RNA, protein and small molecule MTases. They are typically used for enzymatic modification of MTase substrates with unique functional groups which are labeled with reporter groups in a second chemical step. This is exemplified in a protocol for fluorescence labeling of histone H3 protein. A small propargyl group is transferred from the cofactor analogue SeAdoYn to the protein by the histone H3 lysine 4 (H3K4) MTase Set7/9 followed by click labeling of the alkynylated histone H3 with TAMRA azide. MTase-mediated labeling with cofactor analogues is an enabling technology for many exciting applications including identification and functional study of MTase substrates as well as DNA genotyping and methylation detection.

One-step Labeling of DNA via Aziridine Cofactors
This example reaction is carried out with the DNA MTase M.BseCI, which modifies the second adenine residue within the double-stranded 5&#8217;-ATCGAT-3&#8217; sequence and has one recognition site on the pBR322 plasmid (Figure 4A). To test plasmid labeling, pBR322 is challenged with the restriction endonuclease (REase) R.TaqI (5&#8216;-TCGA-3&#8216;). R.TaqI has seven sites on pBR322, one of which is included in the M.BseCI site. If labeling occurs, the M.BseCI site will be protected against cleavage and a new fragment with 683 base pairs (bp) will form while two other fragments will disappear (315 and 368 bp). Of course, when analyzing DNA labeling with other DNA MTases different REases with corresponding recognition sequences should be employed.
The agarose gel in Figure 4B clearly shows the appearance of a new fragment with 683 bp (lane 3). This demonstrates that labeling of the M.BseCI site occurred. Only the assay (lane 3) shows this fragment. The &#8220;cofactor&#8221; control (lane 5) as well as the &#8220;enzyme&#8221; control (lane 7) are missing this band. Especially the &#8220;enzyme&#8221; control is important because it demonstrates that the natural cofactor AdoMet is not present in the enzyme preparation. Specificity of the labeling reaction is demonstrated by electromobility shift assay (EMSA) upon addition of streptavidin (Figure 4B and detail in 4C). Electromobility of the 683 bp is retarded while the mobility of all other fragments without M.BseCI site is unchanged compared to the controls (compare lane 4 with lanes 6 and 8).
<img alt="Figure 4" fo:content-width="7in" src="/files/ftp_upload/52014/52014fig4highres.jpg" width="700" /> 
Figure 4: Sequence specific biotinylation of pBR322 plasmid DNA with M.BseCI and 6BAz. A. Plasmid map of pBR322 showing recognition sites for M.BseCI (red) and R.TaqI (green) as well as the length of expected DNA fragments in base pairs (bp) after restriction with R.TaqI. B. Analysis of DNA fragmentation and EMSA by agarose gel electrophoresis. M = Marker, lanes 1 and 2: plasmid DNA only, lanes 3 and 4: assay, lanes 5 and 6: &#39;cofactor&#39; control, lanes 7 and 8: &#39;enzyme&#39; control. Lanes 2, 4, 6 and 8 additionally contain streptavidin. C. Enlarged inset from B. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52014/52014fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Two-step Labeling of Proteins via Double Activated Cofactors
As an example for two-step labeling of MTase substrates with double-activated AdoMet analogues we chose the histone H3 lysine 4 (H3K4) MTase Set7/9 and the small SeAdoYn cofactor. After enzymatic transfer of the activated propargyl group to the histone H3 substrate (first step), the terminal alkyne is chemically labeled with an azide-modified TAMRA fluorophore by CuAAC click chemistry (second step) (compare Figure 3). Analysis of the labeling reaction is done by SDS-PAGE and in-gel fluorescence detection (Figure 5A). The assay (lane 1) shows a fluorescent band corresponding to histone H3 indicating that the side chain of the cofactor was successfully transferred and the modified protein labeled with the TAMRA fluorophore. No bands are visible in the &#8220;cofactor&#8221; and &#8220;enzyme&#8221; controls (lanes 2 and 3) demonstrating that labeling of histone H3 is mediated by the MTase. Therefore, non-specific chemical labeling either by the cofactor alone (first step) or during CuAAC (second step) can be ruled out. The &#8220;cofactor&#8221; control is performed differently than in the protocol for one-step DNA labeling. This is because precipitation of histone H3 is more efficient in the presence of Set7/9 and omitting the protein MTase can lead to reduced amounts of histone H3 in the loading control. Thus, we added a high concentration of the natural cofactor AdoMet to compete with SeAdoYn. Loading of histone H3 can be easily controlled by staining the gel with Coomassie Blue after fluorescence detection.
MTase substrates can also be labeled with biotin instead of fluorophores by using azide-derivatized biotin in the second chemical step30. Biotinylation of proteins is conveniently analyzed by Western blotting with horseradish peroxidase-conjugated avidin. This is shown in Figure 5B for the biotinylation of histone H3 with Set7/9, SeAdoYn and azide-modified biotin. The assay in lane 1 shows a clear band for the labeled histone H3. The absence of bands in the &#8220;enzyme&#8221; (lane 2) and &#8220;cofactor&#8221; control (lane 3) demonstrates again that labeling is specific for the MTase. The &#8220;cofactor&#8221; control is simply performed in the absence of protein MTase because the Western blot analysis does not require removal of excess biotin by protein precipitation.
<img alt="Figure 5" fo:content-width="7in" src="/files/ftp_upload/52014/52014fig5highres.jpg" width="700" /> 
Figure 5: Labeling of histone H3 with the protein MTase Set7/9 and SeAdoYn followed by click reactions with azide-derivatized labels. A. In-gel fluorescence of TAMRA-labeled histone H3 and controls as well as staining with Coomassie Blue for loading control. B. Western blot analysis of biotinylated histone H3 and controls using avidin-horseradish peroxidase (HRP) conjugate. Experimental conditions were very similar to those for fluorescence labeling in A (enzymatic modification: 6.5 &#181;M histone H3, 10 &#181;M Set7/9, 600 &#181;M SeAdoYn, in 50 mM Tris-HCl, 5 mM MgCl2, 5% glycerol, pH 9.0, 30 &#176;C, 3 hr; chemical labeling: 0.6 mM CuSO4, 0.6 mM THPTA, 50 mM sodium ascorbate, 1.2 mM biotin-azide, 37 &#176;C, 1 hr). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52014/52014fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Sequence-specific Labeling of Nucleic Acids and Proteins with Methyltransferases and Cofactor Analogues at JoVE.com

Sequence-specific Labeling of Nucleic Acids and Proteins with Methyltransferases and Cofactor Analogues

DNA and proteins are sequence-specifically labeled with affinity or fluorescent reporter groups using DNA or protein methyltransferases and synthetic cofactor analogues. Depending on the cofactor specificity of the enzymes, aziridine or double activated cofactor analogues are employed for one- or two-step labeling.

S-Adenosyl-l-methionine (AdoMet or SAM)-dependent methyltransferases (MTase) catalyze the transfer of the activated methyl group from AdoMet to specific ...

sequence-specific-labeling-nucleic-acids-proteins-with

Nanyang Technological University

Research

JoVE Journal

Biology

13.9K Views.  RWTH Aachen University. DNA and proteins are sequence-specifically labeled with affinity or fluorescent reporter groups using DNA or protein methyltransferases and synthetic cofactor analogues. Depending on the cofactor specificity of the enzymes, aziridine or double activated cofactor analogues are employed for one- or two-step labeling.

Video: Sequence-specific Labeling of Nucleic Acids and Proteins with Methyltransferases and Cofactor Analogues

Immunofluorescent Detection of Two Thymidine Analogues (CldU and IdU) in Primary Tissue

Accurate measurement of cell division is a fundamental challenge in experimental biology that becomes increasingly complex when slowly dividing cells are analyzed. Established methods to detect cell division include direct visualization by continuous microscopy in cell culture, dilution of vital dyes such as carboxy&#64258;uorescein di-aetate succinimidyl ester (CFSE), immuno-detection of mitogenic antigens such as ki67 or PCNA, and thymidine analogues. Thymidine analogues can be detected by a variety of methods including radio-detection for tritiated thymidine, immuno-detection for bromo-deoxyuridine (BrdU), chloro-deoxyuridine (CldU) and iodo-deoxyuridine (IdU), and chemical detection for ethinyl-deoxyuridine (EdU). We have derived a strategy to detect sequential incorporation of different thymidine analogues (CldU and IdU) into tissues of adult mice. Our method allows investigators to accurately quantify two successive rounds of cell division. By optimizing immunostaining protocols our approach can detect very low dose thymidine analogues administered via the drinking water, safe to administer to mice for prolonged periods of time. Consequently, our technique can be used to detect cell turnover in very long-lived tissues. Optimal immunofluoresent staining results can be achieved in multiple tissue types, including pancreas, skin, gut, liver, adrenal, testis, ovary, thyroid, lymph node, and brain. We have also applied this technique to identify oncogenic transformation within tissues. We have further applied this technique to determine if transit-amplifying cells contribute to growth or renewal of tissues. In this sense, sequential administration of thymidine analogues represents a novel approach for studying the origins and survival of cells involved in tissue homeostasis.

Immunofluorescent Detection of Two Thymidine Analogues CldU and IdU in Primary Tissue

We have derived a strategy to detect sequential incorporation of thymidine analogues (CldU and IdU) into tissues of adult mice to quantify two successive rounds of cell division. This strategy is useful to detect cell turnover of long-lived tissues, oncogenic transformation, or transit-amplifying cells.

Accurate measurement of cell division is a fundamental challenge in experimental biology that becomes increasingly complex when slowly dividing cells are ...

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry (CCMS)

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity chromatography 1 or Activity Based Protein Profiling 2. Trifunctional Capture Compounds (CCs, Figure 1A) 3 are the basis for a generic approach, in which the initial equilibrium-driven interaction between a small molecule probe (the selectivity function, here S-adenosyl-L-homocysteine, SAH, Figure 1A) and target proteins is irreversibly fixed upon photo-crosslinking between an independent photo-activable reactivity function (here a phenylazide) of the CC and the surface of the target proteins. The sorting function (here biotin) serves to isolate the CC - protein conjugates from complex biological mixtures with the help of a solid phase (here streptavidin magnetic beads). Two configurations of the experiments are possible: "off-bead" 4 or the presently described "on-bead" configuration (Figure 1B). The selectivity function may be virtually any small molecule of interest (substrates, inhibitors, drug molecules).
S-Adenosyl-L-methionine (SAM, Figure 1A) is probably, second to ATP, the most widely used cofactor in nature 5, 6. It is used as the major methyl group donor in all living organisms with the chemical reaction being catalyzed by SAM-dependent methyltransferases (MTases), which methylate DNA 7, RNA 8, proteins 9, or small molecules 10. Given the crucial role of methylation reactions in diverse physiological scenarios (gene regulation, epigenetics, metabolism), the profiling of MTases can be expected to become of similar importance in functional proteomics as the profiling of kinases. Analytical tools for their profiling, however, have not been available. We recently introduced a CC with SAH as selectivity group to fill this technological gap (Figure 1A).
SAH, the product of SAM after methyl transfer, is a known general MTase product inhibitor 11. For this reason and because the natural cofactor SAM is used by further enzymes transferring other parts of the cofactor or initiating radical reactions as well as because of its chemical instability 12, SAH is an ideal selectivity function for a CC to target MTases. Here, we report the utility of the SAH-CC and CCMS by profiling MTases and other SAH-binding proteins from the strain DH5&alpha; of Escherichia coli (E. coli), one of the best-characterized prokaryotes, which has served as the preferred model organism in countless biochemical, biological, and biotechnological studies. Photo-activated crosslinking enhances yield and sensitivity of the experiment, and the specificity can be readily tested for in competition experiments using an excess of free SAH.

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry CCMS

Capture Compounds are trifunctional small molecules to reduce the complexity of the proteome by functional reversible small molecule-protein interaction followed by photo-crosslinking and purification. Here we use a Capture Compound with S-adenosyl-L-homocysteine-binding as selectivity function to isolate methyltransferases from an Escherichia coli whole cell lysate and identify them by MS.

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity ...

NanoDrop Microvolume Quantitation of Nucleic Acids

Biomolecular assays are continually being developed that use progressively smaller amounts of material, often precluding the use of conventional cuvette-based instruments for nucleic acid quantitation for those that can perform microvolume quantitation.
The NanoDrop microvolume sample retention system (Thermo Scientific NanoDrop Products) functions by combining fiber optic technology and natural surface tension properties to capture and retain minute amounts of sample independent of traditional containment apparatus such as cuvettes or capillaries. Furthermore, the system employs shorter path lengths, which result in a broad range of nucleic acid concentration measurements, essentially eliminating the need to perform dilutions. Reducing the volume of sample required for spectroscopic analysis also facilitates the inclusion of additional quality control steps throughout many molecular workflows, increasing efficiency and ultimately leading to greater confidence in downstream results.
The need for high-sensitivity fluorescent analysis of limited mass has also emerged with recent experimental advances. Using the same microvolume sample retention technology, fluorescent measurements may be performed with 2 &mu;L of material, allowing fluorescent assays volume requirements to be significantly reduced. Such microreactions of 10 &mu;L or less are now possible using a dedicated microvolume fluorospectrometer.
Two microvolume nucleic acid quantitation protocols will be demonstrated that use integrated sample retention systems as practical alternatives to traditional cuvette-based protocols. First, a direct A260 absorbance method using a microvolume spectrophotometer is described. This is followed by a demonstration of a fluorescence-based method that enables reduced-volume fluorescence reactions with a microvolume fluorospectrometer. These novel techniques enable the assessment of nucleic acid concentrations ranging from 1 pg/ &mu;L to 15,000 ng/ &mu;L with minimal consumption of sample.

The use of NanoDrop microvolume systems as practical and efficient alternatives to traditional nucleic acid quantitation methodology is described through the demonstration of two microvolume nucleic acid quantitation protocols.

Biomolecular assays are continually being developed that use progressively smaller amounts of material, often precluding the use of conventional ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Lineage Labeling of Zebrafish Cells with Laser Uncagable Fluorescein Dextran

A central problem in developmental biology is to deduce the origin of the myriad cell types present in vertebrates as they arise from undifferentiated precursors. Researchers have employed various methods of lineage labeling, such as DiI labeling1 and pressure injection of traceable enzymes2 to ascertain cell fate at later stages of development in model systems. The first fate maps in zebrafish (Danio rerio) were assembled by iontophoretic injection of fluorescent dyes, such as rhodamine dextran, into single cells in discrete regions of the embryo and tracing the labeled cell's fate over time3-5. While effective, these methods are technically demanding and require specialized equipment not commonly found in zebrafish labs. Recently, photoconvertable fluorescent proteins, such as Eos and Kaede, which irreversibly switch from green to red fluorescence when exposed to ultraviolet light, are seeing increased use in zebrafish6-8. The optical clarity of the zebrafish embryo and the relative ease of transgenesis have made these particularily attractive tools for lineage labeling and to observe the migration of cells in vivo7. Despite their utility, these proteins have some disadvantages compared to dye-mediated lineage labeling methods. The most crucial is the difficulty we have found in obtaining high 3-D resolution during photoconversion of these proteins. In this light, perhaps the best combination of resolution and ease of use for lineage labeling in zebrafish makes use of caged fluorescein dextran, a fluorescent dye that is bound to a quenching group that masks its fluorescence9. The dye can then be "uncaged" (released from the quenching group) within a specific cell using UV light from a laser or mercury lamp, allowing visualization of its fluorescence or immunodetection. Unlike iontophoretic methods, caged fluorescein can be injected with standard injection apparatuses and uncaged with an epifluorescence microscope equipped with a pinhole10. In addition, antibodies against fluorescein detect only the uncaged form, and the epitope survives fixation well11. Finally, caged fluorescein can be activated with very high 3-D resolution, especially if two-photon microscopy is employed 12,13. This protocol describes a method of lineage labeling by caged fluorescein and laser uncaging. Subsequenctly, uncaged fluorescein is detected simultaneously with other epitopes such as GFP by labeling with antibodies.

This protocol delineates a way to label and trace the fate of small groups of cells zebrafish embryos using UV-uncaging of caged fluorescein, followed by whole mount immunolabeling to amplify the signal from the uncaged fluorescein.

A central problem in developmental biology is to deduce the origin of the myriad cell types present in vertebrates as they arise from undifferentiated ...

Concentration Determination of Nucleic Acids and Proteins Using the Micro-volume Bio-spec Nano Spectrophotometer

Nucleic Acid quantitation procedures have advanced significantly in the last three decades. More and more, molecular biologists require consistent small-volume analysis of nucleic acid samples for their experiments. The BioSpec-nano provides a potential solution to the problems of inaccurate, non-reproducible results, inherent in current DNA quantitation methods, via specialized optics and a sensitive PDA detector. The BioSpec-nano also has automated functionality such that mounting, measurement, and cleaning are done by the instrument, thereby eliminating tedious, repetitive, and inconsistent placement of the fiber optic element and manual cleaning.
In this study, data is presented on the quantification of DNA and protein, as well as on measurement reproducibility and accuracy. Automated sample contact and rapid scanning allows measurement in three seconds, resulting in excellent throughput. Data analysis is carried out using the built-in features of the software. The formula used for calculating DNA concentration is:
Sample Concentration = DF &middot; (OD260-OD320)&middot; NACF (1)
Where DF = sample dilution factor and NACF = nucleic acid concentration factor.
The Nucleic Acid concentration factor is set in accordance with the analyte selected1.
Protein concentration results can be expressed as &mu;g/ mL or as moles/L by entering e280 and molecular weight values respectively. When residue values for Tyr, Trp and Cysteine (S-S bond) are entered in the e280Calc tab, the extinction coefficient values are calculated as e280 = 5500 x (Trp residues) + 1490 x (Tyr residues) + 125 x (cysteine S-S bond). The e280 value is used by the software for concentration calculation.
In addition to concentration determination of nucleic acids and protein, the BioSpec-nano can be used as an ultra micro-volume spectrophotometer for many other analytes or as a standard spectrophotometer using 5 mm pathlength cells.

This communication presents data on the accuracy and reproducibility of the BioSpec-nano UV-VIS spectrophotometer for dsDNA and protein quantitation. Even with ultra-small volumes (1 to 2 L), reproducibility is excellent, while the automated wiping function improves throughput and results in minimal carryover for more precise results.

Nucleic Acid quantitation procedures have advanced significantly in the last three decades.  More and more, molecular biologists require consistent ...

ReAsH/FlAsH Labeling and Image Analysis of Tetracysteine Sensor Proteins in Cells

Fluorescent proteins and dyes are essential tools for the study of protein trafficking, localization and function in cells. While fluorescent 
proteins such as green fluorescence protein (GFP) have been extensively used as fusion partners to proteins to track the properties of a protein of interest1, recent 
developments with smaller tags enable new functionalities of proteins to be examined in cells such as conformational change and protein-association 2, 3. One small 
tag system involves a tetracysteine motif (CCXXCC) genetically inserted into a target protein, which binds to biarsenical dyes, ReAsH (red fluorescent) and FlAsH 
(green fluorescent), with high specificity even in live cells 2. The TC/biarsenical dye system offers far less steric constraints to the host protein than 
fluorescent proteins which has enabled several new approaches to measure conformational change and protein-protein interactions 4-7. We recently developed 
a novel application of TC tags as sensors of oligomerization in cells expressing mutant huntingtin, which when mutated aggregates in neurons in Huntington 
disease 7. Huntingtin was tagged with two fluorescent dyes, one a fluorescent protein to track protein location, and the second a TC tag which only 
binds biarsenical dyes in monomers. Hence, changes in colocalization between protein and biarsenical dye reactivity enabled submicroscopic oligomer 
content to be spatially mapped within cells. Here, we describe how to label TC-tagged proteins fused to a fluorescent protein (Cherry, GFP or CFP) 
with FlAsH or ReAsH in live mammalian cells and how to quantify the two color fluorescence (Cherry/FlAsH, CFP/FlAsH or GFP/ReAsH combinations).

The biarsenical dyes FlAsH and ReAsH bind specifically to tetracysteine motifs in proteins and can selectively label proteins in live cells. Recently this labeling strategy has been used to develop sensors for different protein conformations or oligomeric states. We describe the labeling approach and methods to quantitatively analyze binding.

Fluorescent proteins and dyes are essential tools for the study of protein trafficking, localization and function in cells.  While fluorescent 
proteins ...

Metabolic Pathway Confirmation and Discovery Through 13C-labeling of Proteinogenic Amino Acids

Microbes have complex metabolic pathways that can be investigated using biochemistry and functional genomics methods. One important technique to examine cell central metabolism and discover new enzymes is 13C-assisted metabolism analysis 1. This technique is based on isotopic labeling, whereby microbes are fed with a 13C labeled substrates. By tracing the atom transition paths between metabolites in the biochemical network, we can determine functional pathways and discover new enzymes. 
As a complementary method to transcriptomics and proteomics, approaches for isotopomer-assisted analysis of metabolic pathways contain three major steps 2. First, we grow cells with 13C labeled substrates. In this step, the composition of the medium and the selection of labeled substrates are two key factors. To avoid measurement noises from non-labeled carbon in nutrient supplements, a minimal medium with a sole carbon source is required. Further, the choice of a labeled substrate is based on how effectively it will elucidate the pathway being analyzed. Because novel enzymes often involve different reaction stereochemistry or intermediate products, in general, singly labeled carbon substrates are more informative for detection of novel pathways than uniformly labeled ones for detection of novel pathways3, 4. Second, we analyze amino acid labeling patterns using GC-MS. Amino acids are abundant in protein and thus can be obtained from biomass hydrolysis. Amino acids can be derivatized by N-(tert-butyldimethylsilyl)-N-methyltrifluoroacetamide (TBDMS) before GC separation. TBDMS derivatized amino acids can be fragmented by MS and result in different arrays of fragments. Based on the mass to charge (m/z) ratio of fragmented and unfragmented amino acids, we can deduce the possible labeled patterns of the central metabolites that are precursors of the amino acids. Third, we trace 13C carbon transitions in the proposed pathways and, based on the isotopomer data, confirm whether these pathways are active 2. Measurement of amino acids provides isotopic labeling information about eight crucial precursor metabolites in the central metabolism. These metabolic key nodes can reflect the functions of associated central pathways.
13C-assisted metabolism analysis via proteinogenic amino acids can be widely used for functional characterization of poorly-characterized microbial metabolism1. In this protocol, we will use Cyanothece 51142 as the model strain to demonstrate the use of labeled carbon substrates for discovering new enzymatic functions.

Metabolic Pathway Confirmation and Discovery Through 13C-labeling of Proteinogenic Amino Acids

13C-isotope labeling is a useful technique for determining the cell central metabolism for various types of microorganisms. After cells have been cultured with a specific labeled substrate, GC-MS measurement can reveal functional metabolic pathways based on unique labeling patterns in proteinogenic amino acids.

Microbes have complex metabolic pathways that can be investigated using biochemistry and functional genomics methods. One important technique to examine ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

Analyzing and Building Nucleic Acid Structures with 3DNA

The 3DNA software package is a popular and versatile bioinformatics tool with capabilities to analyze, construct, and visualize three-dimensional nucleic acid structures. This article presents detailed protocols for a subset of new and popular features available in 3DNA, applicable to both individual structures and ensembles of related structures. Protocol 1 lists the set of instructions needed to download and install the software. This is followed, in Protocol 2, by the analysis of a nucleic acid structure, including the assignment of base pairs and the determination of rigid-body parameters that describe the structure and, in Protocol 3, by a description of the reconstruction of an atomic model of a structure from its rigid-body parameters. The most recent version of 3DNA, version 2.1, has new features for the analysis and manipulation of ensembles of structures, such as those deduced from nuclear magnetic resonance (NMR) measurements and molecular dynamic (MD) simulations; these features are presented in Protocols 4 and 5. In addition to the 3DNA stand-alone software package, the w3DNA web server, located at <a href="http://w3dna.rutgers.edu" target="_blank">http://w3dna.rutgers.edu</a>, provides a user-friendly interface to selected features of the software. Protocol 6 demonstrates a novel feature of the site for building models of long DNA molecules decorated with bound proteins at user-specified locations.

The 3DNA software package is a popular and versatile bioinformatics tool with capabilities to analyze, construct, and visualize three-dimensional nucleic acid structures. This article presents detailed protocols for a subset of new and popular features available in 3DNA, applicable to both individual structures and ensembles of related structures.

The 3DNA software package is a popular and versatile bioinformatics tool with capabilities to analyze, construct, and visualize three-dimensional nucleic ...

Sequence-specific Labeling of Nucleic Acids and Proteins with Methyltransferases and Cofactor Analogues

Summary

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Chapters in this video

Immunofluorescent Detection of Two Thymidine Analogues (CldU and IdU) in Primary Tissue

Profiling of Methyltransferases and Other S-adenosyl-_L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry (CCMS)

NanoDrop Microvolume Quantitation of Nucleic Acids

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Lineage Labeling of Zebrafish Cells with Laser Uncagable Fluorescein Dextran

Concentration Determination of Nucleic Acids and Proteins Using the Micro-volume Bio-spec Nano Spectrophotometer

ReAsH/FlAsH Labeling and Image Analysis of Tetracysteine Sensor Proteins in Cells

Metabolic Pathway Confirmation and Discovery Through ¹³C-labeling of Proteinogenic Amino Acids

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Analyzing and Building Nucleic Acid Structures with 3DNA